Biofilm ProtocolsA detailed description of the methods used by the 2011

Glasgow iGEM team tomeasure biofilms

Biofilm Protocols

Biofilm formation assay

We were also given tips on forming biofilms by Dr Dan Walker and his PhD student. They told us to keep the following things in mind if we were unable to get hold of a special biofilm-forming device. We then devised our own method of forming biofilms in the lab.

• Biofilm forms thickest at an air-water interface• The media for forming biofilms should be kept in motion throughout the formation process• The slide must not allow planktonic bacteria to rest on it – ie. It must not sit at the bottom

of a container• Biofilm will form on most surfaces, with a few exceptions such as Teflon or other non-stick

materials• Biofilms are not resistant to high level shear forces – they will adhere to each other but not

to the surface they are grown on. Excess shear force will ruin the biofilm structure.

The most efficient method of growing biofilm that we found was to place a glass slide vertically into a 50ml corning tube and fill it with 25ml of media (see figure 1). The media was inoculated with 1μl per ml of over-night culture and relevant antibiotics were added to a desired concentration.

The glass slide was sterilised with 100% ethanol and air-dried in a fume hood to prevent contamination. It is important to observe very sterile technique during this protocol as the is a high risk of contamination.

The lid was then replaced to prevent contamination and the tube was left on a shaker moving at 100 rpm for the desired length of growing time. We grew all of our biofilms at room temperature since we were initially working with Pseudomonas strains which can be hazardous to human health. Once biofilm has grown for the required duration it can either be stained to visualise or fragmented and the number of cells in it counted for a quantitative measurement. These techniques are described overleaf. All experiments were carried out in triplicate to improve accuracy

Figure 1. Glass slide inserted vertically into a 50ml corning tube containing 25 ml of media.

Biofilm gram-staining technique

As biofilms are sensitive to shear-force it is difficult to stain them without causing considerable damage to the structure. As a result, we instead used the following protocol to stain our biofilms with quite a positive effect:

1. Extract the slide from the corning tube slowly and carefully, making sure not to disrupt the biofilm structure where-ever possible

2. Select an area of approximately 2cm2 on the slide to measure biofilm – preferably an area including the air-water interface. Clean the rest of the slide thoroughly with blue-roll and ethanol. Ensure that there is at least 2mm clear on either side of the area you wish to stain.

3. Take a cover-slip and create ridges of vaseline along the edges, to an approximate height of 1mm. These ridges will ensure that the biofilm structure is not crushed when studied under the microscope, they will also allow a complete tunnel for the stains to pass through when coating the biofilm. (See figure 2)

4. Carefully place the coverslip onto the prepared slide, ensuring that there are no gaps along the edges.

5. Run the following reagents across the biofilm by placing drops under it from one side of the coverslip and allowing them to flow underneath until they cover the entire area beneath the slide. When removing these reagents, use a piece of blue-roll to draw the fluid under the slide using capillary force (see figure 3):

1. Add Crystal violet and wait 30 seconds before drawing through.2. Wash through with distilled water3. Add Gram's Iodine and wait 90 seconds before drawing through4. Decolourise with alcohol until no more purple is visible5. Wash through with distilled water6. Counterstain with safrinin and wait 30 seconds7. Wash with distilled water one final time

Figure 2. Push the cover-slip into the vaseline Figure 3. Drop the stain in from one side to create ridges which hold the coverslip away so that it fills the area beneath the cover from the biofilm. slip, then draw it through with blue roll.

Measuring number of cells in a biofilm

When we decided to measure the number of bacteria leaving a biofilm we faced the challenge of creating a measurable method for biofilm dissociation.

• Prepare 2 identical sterile 50ml corning tubes containing 35ml of media. Label these 'biofilm' and 'dissociation'.

• Carefully transfer the biofilm covered slide into the 'dissociation' tube. Leave the slide sitting in this media for 1 hour and then transfer it to the 'biofilm' tube. This step takes into account background dissociation of bacteria leaving the biofilm.

• Using a sterile spatula (washed in ethanol), scrape all sides of the slide to shear biofilm from it into the media.

• Sonicate both the biofilm and dissociation tubes at low power for 30 seconds, to break up the biofilm completely but not lyse the cells.

• Create serial dilutions of these mixtures. The range is specific to how long the bacteria have been forming the biofilm for. We started out using the range undiluted to 10-8

• Plate 200μl of each serial dilution onto agar plates containing the same reagents and leave overnight